CLS  2523   HEMATOLOGY   I

Introduction and Techniques 

This part of the teaching syllabus addresses techniques, safety, and other elements
fundamental to the concepts of basic hematology.  All objectives listed are in the
cognitive domain unless otherwise noted.  The student at the end of the instructional
period, is responsible for meeting these objectives by achieving a cumulative score of
70% or better on all problem sets, case studies, major exams, quizzes, and library
assignments. 

Objectives are listed in numerical order.  The student, upon completion of the
classroom component of this syllabus will be responsible to successfully:

01     BRIEFLY DESCRIBE HOW TO CORRECTLY WASH ONE'S HANDS

[1]   Roll out or have available paper toweling for drying the hands.
[2]   Wet hands (includes wrists) under running water.
[3]   Apply soap and rub thoroughly for up to 15-20 seconds, producing a good lather. 
[4]   Rub completely over the hands and between the fingers.
[5]   Rinse hands and wrists by allowing water to flow from wrists down over fingers.
[6]   Dry hands with a paper towel BEFORE turning off faucets.  Use paper towel to
        turn off the faucet handles. The towel may be used to open the door before
        discarding.

02       LIST SEVEN REASONS FOR WASHING YOUR HANDS

[1]        Between patients.
[2]        After removal of your gloves or between glove changes.
[3]        Before and after the use of the lavatory.
[4]        Before and after drinking, eating, application of cosmetics, smoking, handling
             of contact lenses, etc.
[5]        Any time there is visible contamination of blood or other body fluids.
[6]        Before and after any activity that requires contact with eyes, mucous membranes,
             or injuries in the skin.
[7]        After completion of work. 

03      DESCRIBE SEVERAL WAYS IN WHICH SAFETY CAN BE PRACTICED
IN THE PATIENT’S ROOM

Patient safety is a responsibility for every member of the health care team.  The
following precautions are practical ways that a laboratorian can contribute to
patient safety.
[1]  Proper disposal of specimen collection supplies after collection of the blood
       specimen.
[2]  Note the appearance of the room when you enter:
            A.        Are the bed rails were up or down?   Be sure that the bed
                       rails are as you found them when you leave the room.
            B.         Note the condition of the floors.  If you observe spills of
                       any time, notify the floor personnel of a possible hazard.
            C.        Note the odor of the room.  If it is unusual, notify the
                        nursing desk.
[3]   Observe the patient.  If the patient appears to be in pain or    uncomfortable, notify
        the nursing desk.  It may be helpful to check at the nursing desk and ask about the
        status of the patient.
[4]   Do not touch or handle any equipment around the bed or attached to the patient. 
       Check at the nursing desk if something needs to be moved or adjusted.
[5]   If the patient is receiving IV fluids, note the condition of the IV site.  If it looks
       swollen or out of character, notify the nursing desk. 

04      EXPLAIN THE PURPOSE OR IMPORTANCE OF STANDARD
PRECAUTIONS

Standard precautions consists of a set of prescribed sensible cautions to follow to prevent
the exposure of the laboratory worker to blood borne pathogens.   The application of this
safety practice eliminates the need to have isolation procedures for different types of
infections, the need for warning labels on patient specimens, or separate procedures for
handling the different types of patient specimens.

Standard precautions stipulates such activities as the wearing of gloves, laboratory coats,
eye and face protection, and washing of hands to prevent exposure to pathogens and
transmission of microorganisms.  These precautions describe the proper disposal for
gloves and other potentially contaminated items.   Also are included are rules for
wearing laboratory coats. 

05      EXPLAIN WHY SAFETY IS AN IMPORTANT PRACTICE IN THE HEALTH
CARE SETTING

 All health care providers, who work with patients, are exposed to contagious diseases
spread by blood borne pathogens or other means.  The practice of Standard Precautions
is a part of the laboratory profession that assumes most human blood and body fluids
are infectious (even if they are not).   Serious diseases to guard against are [1] hepatitis A,
[2] hepatitis B, [3] hepatitis C, [4] hepatitis D, [5] hepatitis G, [6] AIDS, and [7] syphilis. 
The most important means of protecting one’s self is the washing of hands (shown to
have a 95% effectiveness in preventing the spread of disease), wearing gloves, wearing
an approved lab coat, and/or wearing safety glasses or face shield.

06    LIST A MINIMUM OF FOURTEEN SAFE PRACTICE RULES FOR THE
LABORATORY
[ 1 ]    Wash hands.
[ 2 ]    Do not eat, drink, smoke, or apply cosmetics in the laboratory.
[ 3 ]    Keep fingers, pens, pencils, and other fomites out of one’s mouth and away from
           the mucus membranes.
[ 4 ]    Do not store food or drink in the lab’s refrigerator.
[ 5 ]    Do not pipet by mouth.
[ 6 ]    Sharps and other needles are to be placed in appropriate containers.
[ 7 ]    Perform procedures that may cause splashing, spraying or otherwise causing
           airborne droplets behind barriers.
[ 8 ]    Wear protective laboratory coats.
[ 9 ]    Wear gloves.
[ 10 ]   Properly label everything
[ 11 ]   Do not wear laboratory coats to cafeteria or break room.
[ 12 ]   Keep fresh bleach for blood and body fluid spills. 
[ 13 ]   Have appropriate warning and caution signs in place.
[ 14 ]   Periodic training in epidemiology and symptoms of blood borne diseases.

 07       LIST EIGHT UNIVERSAL PRECAUTIONS APPROPRIATE FOR THE
 LABORATORY

[1]    Wear gloves when handling any type of body fluid.
[2]    Wear gloves when performing any type of blood collecting procedure.
[3]    Change gloves between patients.
[4]    Use barrier protection if there is a chance of skin or mucous membrane
         exposure to blood or other body fluids.
[5]     Wash hands after removal of gloves or when gloves are known to be
         contaminated or when leaving the laboratory.
[6]     Follow laboratory protocol when handling sharps and disposing of sharps.   Do
         not bend, break, or recap needles.
[7]    Wear aprons over the lab coat if there is a risk of splashing.
[8]    Wear a face shield if there is a risk of splashing.

08      LIST TEN ANTICOAGULANTS THAT MAY BE USED IN THE
LABORATORY AND THEIR PURPOSE

[ 1 ]    Ethylene diaminetetraacetic acid (EDTA): Synonyms:  versene and sequestrene. 
           Comes in a lavender stoppered tube and is used for hematology testing.  Calcium
           is chelated, blocking the coagulation cascade phenomenon.  Commonly used
           anticoagulant in hematology testing.
[ 2 ]    Heparin (Na⁺, NH₄⁺, and Li⁺⁺ salts): Comes in the green stoppered tube.  Used
           primarily for chemistry testing, with the exception of electrolytes.  Inhibits
           coagulation by interfering with thrombin.
[ 3 ]    Heparin (Li⁺⁺ salts): Is marketed in a green and grey marbled stoppered tube. 
          Used primarily for electrolyte testing.  It is appropriate for other chemistry test
           procedures.  Inhibits coagulation by interfering with thrombin.  NOTE: Some
           manufacturers are marketing this tube with a light green closure.
[ 4 ]    Sodium citrate: Comes in a light blue stoppered tube as a liquid.  It chelates
           calcium, blocking the coagulation cascade phenomenon.  It principle use is in
           coagulation testing.  If there is platelet satellite phenomenon, collect a blood
           specimen in this anticoagulant for retesting.
[ 5 ]    Potassium oxalate with sodium fluoride: Identified in the grey stoppered tube. 
           The potassium oxalate prevents clotting by chelating calcium and the sodium
           fluoride inhibits glycolysis.  This tube is used in glucose and alcohol testing.
[ 6 ]    No anticoagulant: The red stoppered tube may contain an internal silicone
           coating to facilitate uniform clotting.  Used in chemistry, blood bank, and
           serology. 
[ 7 ]    Heparin (Na⁺ salt): Marketed in a royal blue stoppered tube.  It is used in testing
          for trace elements.  It inhibits coagulation by interfering with thrombin
[ 8 ]    Sodium polyanetholsulfonate (SPS): A yellow stoppered tube containing a liquid
           anticoagulant.  It blocks coagulation and neutralizes the antibacterial properties
           of serum, inhibits complement activity, inhibits phagocytosis, and inactivates
           aminoglycosides.  Its principle use is in microbiology, but has been employed in
           the blood bank.
[ 9 ]    No anticoagulant: This tube has a red and gray marbled stopper.  The tube
           contains silica particles to augment the clotting phenomenon.  Its primary use is
           in chemistry.   NOTE: Some manufacturers are marketing this tube with a gold closure.
[ 10 ]    Thrombin: A tube with a yellow and gray marbled stopper.  Thrombin
             accelerates the clotting process.  Used primarily for STAT chemistries.
             NOTE: Some manufacturers are marketing this tube with an orange closure.

The most commonly used anticoagulants in hematology are:
[ 1 ]    EDTA for complete blood counts.
[ 2 ]    Sodium citrate for coagulation studies. 
NOTE:  Some hematology procedures require heparin (example: osmotic
                fragility testing). 

09  EXPLAIN WHAT CLOT ACTIVATORS ARE AND WHY THEY ARE USED

Clot activator are any substance that will initiate and accelerate clotting, shortening the
time required for coagulation.  They are found in the bottom of the tube as an off-
colored substance or they may coat the inside walls of the test tube.  Specific types of
activators are [ 1 ] silica particles and [ 2 ] thrombin.

10   EXPLAIN WHAT SEPARATOR GELS ARE AND WHY THEY ARE USED

This is an inert gel substance that can change it viscosity under centrifugal pressure.  It is
designated as a thixotropic substance and will force itself upward against the centrifugal
pressure as the cellular element press toward the bottom of the tube.  Serum is carried
upward in front of the gel.  When the centrifugation process is completed, there will be
a gel barrier between the red blood cells and serum. This technology allows for
immediate separation of serum from cellular elements for immediate testing.  It is
permissible to leave the serum in the tube (on top of the gel barrier) for up to 48 hours.
CAUTION:   Do not use this blood for blood banking procedures.

11        LIST 26 STEPS FOR A CORRECT VENIPUNCTURE IN AN ADULT
PATIENT

[ 01 ]        Prepare the accession order (this may have been already performed).
[ 02 ]        Identify the patient.
[ 03 ]        Verify any patient diet restrictions.|
[ 04 ]        Assemble supplies for the venipuncture and put on gloves.
[ 05 ]        Reassure the patient.
[ 06 ]        Position the patient if necessary.
[ 07 ]        Verify the paperwork and blood collecting tubes.
[ 08 ]        Ensure that the patient’s hand is closed.
[ 09 ]        Select the venipuncture site.     
[ 10 ]      Clean the venipuncture site.
[ 11 ]      Place the tourniquet 3-4 inches above the venipuncture site.
[ 12 ]      Inspect the needle (and other items as necessary).
[ 13 ]      Perform the venipuncture.
[ 14 ]      Mix those tubes with additives by gentle inversion as each tube is collected.
[ 15 ]      Release the tourniquet and remove it.
[ 16 ]      Tell the patient to relax and open their hand.  Reassure if necessary.
[ 17 ]      Place a gauze or cotton ball over the puncture site, then remove the needle as
               you firmly position the gauze or cotton ball over the wound.  Allow the
               patient to hold it firmly in place with the other hand.
[ 18 ]      Remove last tube from vacutainer needle.
[ 19 ]      Replace needle sheath and discard in biohazard container.
[ 20 ]     NOTE: If a syringe was used instead of a vacutainer, fill the tubes, mixing each
              tubes after it is filled.  Discard the syringe and needle in a biohazard container. 
[ 21 ]     Place a bandage over the wound.  (If the patient declines, omit the bandage
              and do not make an issue of it.)
[ 22 ]     Some specimens required being maintained at a cold temperature.  Ensure this
              step before leaving the bedside.
[ 23 ]     Advise the proper personnel that the specimen has been collected and diet
              restrictions, if any, may be removed.
[ 24 ]     Label the requisition with the time collected and your initials.
[ 25 ]     If the tubes were not labeled prior to the venipuncture, label the tubes at the
              bedside of the patient.
[ 26 ]     Return the appropriately labeled tube to the correct laboratory area for testing.    

12        EXPLAIN WHAT IS MEANT BY ACCESSION ORDER

This refers to the requisition that has been issued for the patient authorizing testing
and/or treatment.  Hospitals and clinics use computers to assign a work order number,
called an accession number or order, to each requisition.  Preprinted specimen labels with
bar-codes may also be issued.  These accession numbers for the patients specimens and
requisitions must be identical.  It is good practice to re-verify these number before
and after.

 13        LIST ELEVEN REASONS FOR REJECTING A BLOOD SPECIMEN

[ 01 ]      The requisition and label do not match.
[ 02 ]       The tube is unlabeled.
[ 03 ]      The I.D. number is incorrect.
[ 04 ]      The specimen is hemolyzed.
[ 05 ]      The specimen was collected at the wrong time.
[ 06 ]      The specimen was collected in the wrong tube.
[ 07 ]      The specimen contains clots in a tube with an anticoagulant.
[ 08 ]      The specimen is lipemic.
[ 09 ]      If the blood was collected without dietary restrictions.
[ 10 ]      Slow transport of the specimen to the laboratory.
[ 11 ]      Failure to record time and date of collection.

14      LIST EIGHTEEN SOURCES OF ERROR IN PHLEBOTOMY COLLECTIONS

[ 01 ]      Improper patient identification.
[ 02 ]      Reassuring the patient to avoid stress.
[ 03 ]      Failure to verify any diet restrictions.
[ 04 ]      Use of wrong blood collection tube.
[ 05 ]      Improper cleansing of venipuncture site.
[ 06 ]      Inserting needle bevel side down.
[ 07 ]      Using a small gauge needle and inducing hemolysis.
[ 08 ]      Performing the venipuncture in the wrong place (example: above an IV site)
[ 09 ]      Leaving tourniquet on during venipuncture process.
[ 10 ]      Failure to mix blood in tubes containing additives.
[ 11 ]      If using a syringe, drawing on the plunger forcefully and inducing hemolysis.
[ 12 ]      Releasing the tourniquet after withdrawal of the needle.
[ 13 ]      Not applying pressure to the venipuncture site and a hematoma develops.
[ 14 ]      Vigorous shaking of tubes to mix anticoagulants.
[ 15 ]      If using a syringe, when transferring the blood to tubes, applying force to the
              plunger to fill the vacutainer tube faster.
[ 16 ]      Mislabeling tubes.
[ 17 ]      Failure to record time, date, and phlebotomist’s initials.
[ 18 ]      Slow transport to the laboratory.

 15        EXPLAIN WHAT TO DO IN THE EVENT OF AN ADVERSE REACTION

The two most common adverse reactions are syncope and hematomas.  In the event of
beginning syncope (the patient indicates that they feel faint), stop the phlebotomy
procedure immediately, and quickly removing the needle and releasing the tourniquet. 
If the patient in sitting, lower the patient’s head, have the patient take deep breaths,
and apply cool wet cloths/compresses to the back of the neck.  A drink of cold water is
often helpful.  If the patient faints and collapses, lower the patient to the floor to a
supine position.  Apply cold compresses.  The patient will recover, feeling foolish and
somewhat embarrassed.  Reassure the patient that this is not an uncommon happening
and that all is well. 

The second adverse reaction is the sudden development of a hematoma.  This is the
result of  a substantial amount of blood leaking into the surrounding tissues from
around the needle (which may be due to the bevel of the needle protruding partially
from the vein.  If swelling is observed around the area of the needle, stop the
phlebotomy procedure and remove the needle and tourniquet.  Apply a cotton ball
or gauze pad as the needle is withdrawn and apply a firm pressure to the site for a
minimum of two minutes.  Caution: If a firm pressure is not applied to the
venipuncture site or held for an appropriate amount of time after the collection
of blood, a hematoma may develop.

16    LIST SIX REASONS WHY BLOOD MAY NOT BE DRAWN DURING A
VEINPUNCTURE

[ 01 ]    The needle was inserted through the vein.
[ 02 ]    The needle was partially inserted .
[ 03 ]    The bevel of the needle was inserted in a way that it rests against the wall of
             a vein.
[ 04 ]    The needle was inserted too close to a valve that blocks blood flow.
[ 05 ]    The vein has collapsed.
[ 06 ]    The vacutainer tube may contain a partial vacuum or no vacuum.

17        DESCRIBE HOW BLOOD SHOULD BE COLLECTED DURING IV THERAPY

The general rule is that blood is to never be drawn from an arm in which an intra-
venous (IV) catheter has been inserted.  If there is no choice but to draw blood from
such a site, the following is recommended.  [ 1 ] Request that the nurse stop the IV for
at least two minutes.  [ 2 ] Place the tourniquet below the IV site.  [ 3 ] Draw blood
from below the IV site.  [ 4 ] Record on the requisition that the blood was drawn from
the arm that had an IV therapy set up.   

18      EXPLAIN WHY THE TOURNIQUET SHOULD BE RELEASED AFTER
SUCCESSFULLY INSERTING THE NEEDLE INTO THE VEIN

To avoid the problem of hemoconcentration.  Hemoconcentration occurs when the
tourniquet remains on during the time of blood collection.   It is recommended that
the tourniquet should not remain on more than one minute.  After this time the
tourniquet should be released and the patient’s arm “rest” for about three minutes. 
The tourniquet can be reapplied and the venipuncture performed.   There are
exceptions to this rule.  Ignore this rule if the patient has veins that are “fragile”,
tending to collapse or where removal of the tourniquet may cause stoppage of blood
flow.  If the venipuncture is performed smoothly and quickly, then it is possible to
collect the blood specimens in less than one minute time limit allowing the tourniquet
to remain on the arm before releasing it.

 19        LIST FIVE CONSIDERATIONS FOR LOCATING SITE OF THE
VENIPUNCTURE

[ 1 ]   The median cubital vein is preferred because it is large, close to the surface of the
          skin, and sufficiently anchored to the tissue for a successful venipuncture.
[ 2 ]    Avoid the cephalic, median, and basilic veins because of their tendency to
           bruise easily.
[ 3 ]    If the inner aspect of the elbow is not a good site for a venipuncture, consider
          alternate sites such as the ventral forearm, back of the hand, wrist area, ankle,
          and foot.  CAUTION: If the lower extremities are selected as a possible
           venipuncture site, check with the nurse (or physician) to be sure the patient does
           not have extremity problems if they are a diabetic or have a hemoglobinopathy.
[ 4 ]    Avoid areas with scars, hematomas, burns, or edema.
[ 5 ]    Avoid sites with an IV catheter, receiving therapy.
        

 20      LIST THREE COURSES OF ACTION TO FOLLOW IF IT IS DIFFICULT
 TO IDENTIFY A PROMINENT VEIN

[ 1 ]   Check and see which is the dominant arm.  The most prominent veins are located
          in the dominant arm.
[ 2 ]    It is okay to close your eyes while palpating for the vein.  This simple actions
          tends to augment the sense of touch.
[ 3 ]    Have the patient slowly open and close his fist.  Avoid rapid pumping of the fist
           as it tend to cause hemoconcentration.
[ 4 ]    Check the tightness of the tourniquet.  If a tourniquet is too tight, it may occlude
           the arteries, resulting in the failure of the blood to flow into the veins and
           increasing venous pressure.  If the arm turns shades of red or purple, the
           tourniquet is too tight.
[ 5 ]    Massaging the arm may help the veins to engorge and stand out.

 21        EXPLAIN WHY THE BLOOD SAMPLE MUST BE DILUTED BEFORE
TESTING

 Blood is a concentrated solution and must be diluted for enumeration for most tests,
whether performing manual or automated procedures.  Remember that blood, once
diluted becomes unstable and it keeping time is variable, dependent upon the diluent. 
Platelets are very fragile and once diluted must be counted/tested quickly.  Do not
perform platelet counts after 30 minutes.  RBC’s and WBC’s are somewhat hardy
and diluted samples may produce reliable results for up to two hours.   Remember
that evaporation changes the concentration and alters results.

 22        EXPLAIN WHY SALINE (0.85% OR 0.9% NaCl) IS A SATISFACTORY
DILUENT

 Saline is iso-osmotic with cellular elements.  It can be used with cellular counts and
preparing cell suspensions.  Cell suspensions are seldom used in hematology but are a
usual procedure in blood banking.  Diluted blood specimens should be discarded after
about two hours.

 23      BRIEFLY DESCRIBE THE PURPOSE OF EACH OF THE FOLLOWING
DILUTING SYSTEMS 

AMMONIUM OXALATE SOLUTION (1%).  May be used for platelet counts.  It consists
of ammonium oxalate and distilled water.  It could be used also as a WBC counting
solution for hemocytometers.
DRABKIN’S SOLUTION.  Designed for hemoglobin determinations.  It contains
sodium bicarbonate, potassium cyanide, potassium ferricyanide, and distilled water.
MANNER’S DILUTING FLUID.  Designed for eosinophil counts.  It contains phloxine,
trisodium citrate, and distilled water.
PILOT’S SOLUTION.  Designed for eosinophil counts.  It contain propylene glycol,
distilled water, and phloxine.
REE’S AND ECKER SOLUTION.  Designed for the platelet count.  It contains brilliant
cresyl blue, sodium citrate, formalin, and distilled water.

24     BRIEFLY DESCRIBE HOW TO PREPARE A BLOOD SMEAR

[ 01 ]    A drop of blood is placed on one end of a 3" by 1" glass slide and using a
            spreader slide at an angle of approximately 25^o to 45^o, a wedge type smear is
            made and allowed to dry. 
[ 02]    Smears may be made using two cover glasses.  A drop of blood is placed in the
           center of  one of the cover glasses.  The other cover glass is placed over the drop
           of blood so that the corners of each cover glass form an eight-pointed star.  The
           two cover glasses are pulled apart and allowed to dry face up. 

The slides, if not to be immediately stained, must be fixed within two hours to prevent
distortion and deterioration. Methanol is the preferred fixative.  Once stained, if a slide
is to be retained, should be “cover-slipped” to prevent stain deterioration.  Color will
begin to fade in about 2 years on an “uncovered” slide.  Refer to your lab manual for
a more detailed description.

25        LIST SIX COMMON CAUSES FOR A POOR WEDGE-TYPE SMEAR

Common failures encountered in prepare blood smears include [ 1 ] using to large or
too small a drop of blood, [ 2 ] moving the spreader slide in a jerky manner, [ 3 ] failure
to hold the spreader slide firmly on the slide during the “sliding” process, [ 4 ] holding
the spreader slide at the wrong angle, [ 5 ] failure to push the spreader slide to the end
of slide, and [ 6 ] moving the spreader slide to fast or to slow, which affects the
thickness of the smear.

26        DESCRIBE HOW TO MAKE A 2% CELL SUSPENSION

Any size tube will work for this procedure.  Use two drops of fresh blood (normal
hematocrit and hemoglobin) per 4.0 mLs of diluent. Mix the cells and diluent by
inverting several times.  Do not shake to mix.  To be more exact, 0.4 mLs of
blood may be added to 10 mLs of diluent (0.85% or 0.9% NaCl).

 27        LIST EIGHT CONCERNS IN PREPARING A COVERSLIP BLOOD FILM

[01]   The drop of blood must not be large.  A large drop of blood creates a thick smear.
[02]   The drop of blood must not be too small as it may create too thin a film or a
         tiny examination area.
[03]    Watch the timing in pulling the coverslips apart.  Delaying the separation may
         result in the cover-slips sticking to each other resulting in platelet clumping and
         uneven distribution of WBC’s.
[04]    If the coverslips are pulled apart too quickly, an uneven film or even a thick film
        may result.
[05]    Maintain an even horizontal pull.  An uneven separation may create an uneven
        distribution of cells and/or an uneven film.
[06]    The coverslip surface is small and this may interfere with mixing the stain and
         buffer or the rinse process.
[07]    Keep coverslips in pairs, do not intermix.
[08]    Air dry as quickly as possible.

28        LIST FOURTEEN CONCERNS WITH THE WEDGE TYPE BLOOD SMEAR

The wedge-type blood smear is the most widely procedure for blood films.  Concerns are:
[01]   A tendency for poor distribution of nucleated cells.  Monocytes and neutrophils
          tend to accrue in the feathered end of the wedge smear, leaving the central
          examination area deficient.
[02]   Lymphocyte differential counts may be artificially increased because of the
          tendency of monocytes and neutrophils to appear in the feathered edge.
[03]   Trauma to cells is greater in the wedge preparation increasing the number of
          basket or smudge cells.  If this is a problem, add 2--3 drops of 22% bovine
          albumin to 1.0 mL of blood, mix, and make a new smear.
[04]   Large cells then to settle in the edges, which leaves abnormal cells in the center
         and easier to find.
[05]   Too large of a drop of blood will produce a thicker smear.  This causes nucleated
          cells to shrink and stain very intensely.  Red blood cells tend to form rouleaux’s
          so that they cannot be evaluated.
[06]   A large angle on the spreader slide causes a thick smear.
[07]   Spreading the film too fast promotes a thick smear.
[08]   Too small of a drop of blood usually results in a blood film that is too small
          and/or thin.  This results in more spheroid shaped RBC’s, distorted RBC shapes,
          increased numbers of smudge cells, and more nucleated cells accumulate on the
          periphery of the smear.
[09]   Spreading the film too slow promotes a thin smear.
[10]  Using a decreased spreading angle causes thinner blood films.
[11]  Using a spreader slide with rough or dirty edges tend to produce blood films with
         jagged and/or gritty ends.  Gritty areas are usually characterized by increased
         numbers of WBC’s.
[12]  Appearance of “tailed” areas which will demonstrated abnormal increases
         of WBC’s.
[13]  Delay in spreading the blood drop after it has been placed on the slide causes
         grittiness or appearance of tails.
[14]  Using heparinized blood. This may cause nucleated cells to accumulate in the
         thin, tail end.  Heparin also causes formation of a bluish background in the blood
         film when stained.

 29        DESCRIBE THE APPEARANCE OF A GOOD QUALITY WEDGE-
SHAPED SMEAR

 Look for a thick band at the application point.  The cells will be stacked, overlapping,
and closely spaced.  The WBC’s tend to be of small size in this area.  In the thin and
feathery portion, an increase in artifacts will be observed due to the wide separation of
cells.  Wide spaces are characteristic between the cells.  The cells tend to be thinner, with
a larger sized appearance.  Do not evaluate RBC morphology in the thick and thin
areas.  The intermediate region of the slide is characterized by cells that do not touch or
almost do so.  The monocytes and granulocytes tend to be settled out in the “sliding”
process near the lateral edges of the slide.  They can also be noted in larger numbers in
the feathery area of the slide.  Lymphocytes tend to randomly distribute on the slide. 
Other factors to look for are [ 1 ] absence of waves, holes, and ridges and [ 2 ] a smooth
and even appearance.

 30        LIST THREE ADVANTAGES OF THE WEDGE-SHAPED SMEAR

 This type smear is easy to make, requiring less time.  There is less glass breakage than
with the coverglass method because of the ease in handling.  It is the easiest of the
methods to learn.  It is easier to label this slide.  The slide is easy to store and does
not require mounting with permount media and a coverglass.

 31        LIST FIVE DISADVANTAGES OF THE WEDGE-SHAPED SMEAR

 Good quality smears are not the rule.  Disadvantages are as follows: [ 1 ] it is not
possible to produce a uniform slide, [ 2 ] quality affected by hematocrit, [ 3 ] quality
affected by viscosity, [ 4 ] WBC’s are not randomly distributed, [ 5 ] RBC distortion
occurs on the periphery of the smear.

 32        DESCRIBE A BUFFY COAT AND HOW TO PREPARE A BUFFY
COAT SMEAR

 The buffy coat preparation may be required if the following are suspected:
[1]   a blood specimen that is pancytopenic and the abnormal, immature, or reactive
        cell densities are low,
[2]   examining a patient diagnosed with megaloblastic anemia for nucleated red
        blood cells and/or hypersegmented neutrophils,
[3]    looking for plasma cells,
[4]    tumor cells in blood indicating metastasis,
[5]    facilitate the search for bacteria and/or parasites (NOTE: erythrocytes containing
        malarial parasites tend to concentrate at the top of the red cell layer).

The buffy coat is prepared by filling a Wintrobe tube or hematocrit tube with blood. 
It is recommended that the Wintrobe tube be centrifuged at 1000 rpm for six minutes. 
The hematocrit tube for a shorter time in the hemotocrit centrifuge, 2 minutes is
recommended.  These reduced centrifuge times will cause less cellular distortion.  For
the hematocrit tube, score the tube just below the red cell line and break.  Touch the
tube with the buffy coat to a glass slide and allow a small amount of plasma to add to
the buffy coat.  Mix the buffy coat in the plasma and then make the buffy coat film, air
dry and stain.   The buffy coat in the Wintrobe may be aspirated with a pipet and
transferred to a watch glass and mixed with plasma.  Make the buffy coat film, the
air dry and stain.

33        DISCUSS WRIGHT’S STAIN AND ITS ROLE IN STAINING BLOOD FILMS

Wright’s stain is composed of oxidized methylene blue and eosin azures.  It is a
commonly used modification of the Romanowsky stains.  Wright’s stain is made up in
absolute methanol (serves as a fixative) to be a solution of an acid dye (eosin) and a basic
dye (methylene blue).  The quantity of dyes used in making up Wright’s stain is designed
to produce a neutral compound.  The basic dyes in this stain have an affinity for the
acidic components in the cells (nucleus and some cytoplasmic structures) imparting a
violet-blue color.  The “azures” will impart red-purple coloration, augmenting the
polychrome nature of the stain.  The acidic dyes have an affinity for the basic
components, hemoglobin and eosinophilic granules, imparting a orange-red color.
Neutrophil granules contain a slightly predominate amount of acidic substances which
will stain weakly with the azure component of the dye.  The polychrome nature of
Wright’s stain produces a complex staining pattern that will facilitate visual
identification of most cells in circulation.   In the staining process, Wright’s stain is
added to cover the slide.  During this one to two minute interval, the blood film is
being fixed by the alcohol.  An equal amount of  buffer (a pH of 6.4 is recommended) is
added to change the ionization characteristics of the stain and it is during this interval
that the blood film is stained.   Eosin, being acidic, carries a negative charge which
attaches to positive charged molecules.  Methylene blue and its azures carry a positive
charge which binds to negative charged molecules.  The stain and buffer are well mixed
when a green metallic sheen can be seen on the top of the staining mixture.  After
allowing the slide to stain from 30 seconds to 3½ minutes the buffer-stain is flooded
off with distilled water to remove excess coloration and any precipitated stain. 

 Because each batch of Wright’s stain will vary, new staining conditions must be re-
established. Wright’s stain continues to undergo molecular conversions and over time
the staining characteristics may change, requiring that new staining conditions be
established again. 

34        DESCRIBE THE GIEMSA STAIN

 The Giemsa stain does not employ the use of sodium carbonate to oxidize the
methylene blue to produce the methylene azures.   Acid bichromate is used instead to
form a “converted” azure compounds.  Alone, this stain is not a satisfactory stain for
RBC’s, platelets, or WBC cytoplasm.   When used in combination with Wright’s stain,
it can intensify the nuclear features as-well-as the neutrophilic granules and toxic
granules found in the WBC cytoplasm.

25        BRIEFLY DISCUSS ROMANOWSKY STAINS

Romanovsky stain is a polychrome stain that has undergone many modifications
since its discovery by the Russian physician Romanowsky.  These stains are composed
of methylene blue, oxidative products of methylene blue (known as methylene azures
or simply azures), and eosin dyes.  These dyes are not water soluble but will dissolve
in absolute methanol.  Popular modifications of these dyes are Wright’s stain, Giemsa
stain, Jenner’s stain, May-Grunwald stain, and May-Grunwald-Giemsa stain.  There is
little difference in the staining properties of these stains. 

36      DESCRIBE THE APPEARANCE OF A PROPERLY STAINED BLOOD FILM
WITH  WRIGHT’S STAIN

The macroscopic appearance will have a pinkish to pinkish-blue tone.  The
microscopic view will show the following:
[ 01 ]    RBC’s have a pink to orange color,
[ 02 ]    reticulocytes take on a grey-pink to bluish color,
[ 03 ]    lymphocytes have a dark purple or blue nucleus and the cytoplasm is sky blue
             to medium blue,
[ 04 ]    the neutrophil’s nucleus stains dark purple or blue and the granules in the
             cytoplasm are lilac or pink to violet,
[ 05 ]    the nucleus of the monocyte is light purple to grey-blue and the cytoplasm is
             grey-blue with fine red granules,
[ 06 ]    the nucleus of the eosinophil stains like that of the neutrophil, but the large
            cytoplasmic granules stain orange-red,
[ 07 ]    the basophil nucleus stains like that of the neutrophil, but the large cytoplasmic
             granules stain dark blue-black,
[ 08 ]    platelets take on a light blue to purple stain with violet to purple granules.

37       DISCUSS AND LIST FIVE CAUSES FOR A WRIGHT’S STAIN THAT IS
EXCESSIVELY BLUE

[ 01 ]        The staining time may be too long,
[ 02 ]        the stain may be too alkaline,
[ 03 ]        the buffer may be too alkaline,
[ 04 ]        the rinsing technique was incomplete,
[ 05 ]        the smear may be too thick. 

Visual clues to an alkaline stained slide are [A] eosinophil granules are pale and appear
gray, [B] RBC’s appear green, and [C] neutrophil granules appear larger than usual.

38        DESCRIBE FOUR CAUSES FOR A WRIGHT’S STAIN THAT IS
OVERLY RED

[ 01 ]        The staining time is too short.
[ 02 ]        The stain was over rinsed.
[ 03 ]        The stain may be too acid
[ 04 ]        The buffer may be too acid.

Visual clues to an acidic stain are: [A] RBC’s appear bright orange-red, [B] leukocyte nuclei appear light blue, [C] eosinophil granules appear a bright red

39        EXPLAIN HOW THE BUFFER SOLUTION ON WRIGHT’S STAIN WORKS

 Buffer adjusts the pH of the stain and controls the amount of acidic and basic dye that binds to the cell parts.  If the buffer adjusts the pH downward (acid side), more acid dye is taken up, increasing the intensity of the red coloration.  Less basic dye is taken up, therefore less blue coloration.  If the buffer adjusts the pH upward (alkaline side), less red stain is taken up and more blue stain is absorbed producing excessively blue colors.

 40        LIST SIX REASONS WHY PRECIPITATE MAY APPEAR ON A
STAINED BLOOD FILM

[ 1 ]    Insufficient washing/rinsing,
[ 2 ]    stain dried out on the slide,
[ 3 ]    dirty slide,
[ 4 ]    slide may not have lain flat/horizonal on the staining rack,
[ 5 ]    stain may need to be filtered,
[ 6 ]     dust is settling on the slide.

41        EXPLAIN WHAT HAPPENS WHEN OLD BLOOD IS USED TO MAKE
BLOOD FILMS

Old blood behaves in a sludgy manner.  It is best not to use blood that is more than
two hours old.  The following occurs with slides prepared from old blood:
[ 01 ]   uneven distribution of WBC’s,
[ 02 ]   increased number of basket and smudge cells,
[ 03 ]   distortion of cells due to anticoagulant action,
[ 04 ]   rouleaux formation,
[ 05 ]   increased clumping of platelets,
[ 06 ]   the larger leukocytes accumulate along the edge of the slide,
[ 07 ]   increase in artifact formation.

42        BRIEFLY DISCUSS THE PEROXIDASE STAIN AND HOW IT IS PERFORMED

Synonym: Myeloperoxidase stain.  This is a reaction that is dependent upon the
presence of peroxidase in the primary granules of neutrophils, eosinophils, and
monocytes.  Lymphocytes and basophils, and erythrocytes do not have such activity. 
Peroxidase activity is found in the promyelocyte to neutrophil stages. The level of
peroxidase increases as the cells mature.  The neutrophil has the greatest activity,
followed by the eosinophil.  The monocyte’s peroxidase activity is limited to its small
and fine granules.  This stain will help differentiate acute myleogenous leukemia  (FAB
subclasses M1, M2, and M3) and and acute monocytic leukemia (FAB subclass M5)
from acute lymphocytic leukemias (FAB subclasses L1 - L3).  NOTE: The peroxidase in
these cells should be designated as myeloperoxidase (MPO) to distinguish it form other
peroxidases.  The procedure requires preparing smears of blood and/or bone marrow,
air drying and fixing in a special buffer.  The slides are incubated in a solution
containing the substrate, 3-amino-9-ethyl carbazole and hydrogen peroxide.  The
slides are counterstained with hematoxylin.  The presence of reddish-brown to blue-
black granules indicates the presence of the peroxidases, a positive stain.  Interpretation
is as follows:
[ 01 ]   Look for positive activity in the more mature cells.  Generally, the reactivity in
            the mature cells (bands and 'segs') is not significant in differentiating acute
            leukemias.
[ 02 ]   Monocyte peroxidase activity will be slight (weakly positive).  In acute
            monocytic leukemia, the monoblasts are usually negative.  The positive
            reactions are noted in the more mature monocytes.
[ 03 ]   Auer rods are strongly peroxidase positive.  This stain readily demonstrates
            their presence.
[ 04 ]    Eosinophilic granules are strongly peroxidase positive.
[ 05 ]   The following cells are usually peroxidase negative: 
            A.    early myeloblasts
            B.    erythroblasts
            C.    lymphocytes
            D.    mature basophils
            E.    plasma cells

 43        BRIEFLY DISCUSS THE PRUSSIAN BLUE STAIN AND HOW IT IS
PERFORMED.

 Synonyms: Siderocyte stain or Prussian blue reaction.  Free iron (Fe⁺³ state) is seen
as small blue or blue-green siderocyte granules found in the cytoplasm of developing
RBC’s.  This is iron that has not been incorporated into hemoglobin.  If one or more
of these free iron granules are observed, the RBC is called a siderocyte.  In the healthy
individual, up to 1% of the RBC’s may be siderocytes.  Increased siderocytes are
observed in thalassemia major, lead poisoning, leukemia, alcoholism, hemolytic
anemias, megaloblastic anemias, and splenectomy.  Increased siderocytes are an
indicator of abnormal hemoglobin synthesis. Siderocyte granules are observed in
nucleated red blood cells and may be found in the bone marrow reticulocytes.  These
granules are not normal to the mature erythrocytes seen in peripheral blood.    Blood
slides are air dried, then fixed in methanol.  The slides are stained in Prussian blue
reagent (sometimes called Perl’s reagent) for up to 30 minutes.  The slides are then
counterstained with eosin or safranin.  To determine the percentage of siderocytes,
do this: FIRST:  determine the number of siderocytes per 1,000 RBC., SECOND:
use the formula # siderocytes counted / 1000 RBC’s (times) 100, and THIRD:
report the percent siderocytes.  Sample problem: 65 siderocytes divided (÷) by 1000
RBC then the answer is multtiplied (×) 100 = 6.5%

44    BRIEFLY DISCUSS THE SUDAN BLACK B STAIN AND HOW IT
IS PERFORMED

 Sudan black B (SBB) stain is a fat-soluble dye that stains intracellular lipids (sterols,
phospholipids, and neutral fats).  This stain parallels the peroxidase stain results and
it is used to differentiate acute myelogenous and myelomonocytic leukemias from
the acute lymphocytic leukemias.  Those cells that demonstrate MPO activity tend
to demonstrate sudanophilia. This stain, if used, will be employed to differentiate
blast cells in the FAB subclasses (M1 [acute myelocytic leukemia, without maturation],
M2 [acute myelocytic leukemia, with maturation), and M3 [acute promyelocytic
leukemia]) from acute lymphocytic leukemia (ALL).  When interpretating this stain,
 the presence of brown-black granules is considered as a positive stain.  Those cells in
the myelocytic series stains most strongly.  Eosinophils will stain strongly and
monocytes stain weakly as in the peroxidase stain.  The early myeloblasts,
erythroblasts, lymphoblasts, lymphocytes,  megakaryoblasts, platelets, and mature
basophils do not stain.  There is an unusual exception, patients with Burkitt’s
lymphoma, the immature vacuolated lymphocytic appearing cells may stain positive. 
It has been observed that a patient diagnosed with chronic lymphocytic leukemia, in a
blast crisis, may demonstrate positive SBB stain.

 Observations regarding SBB are: [ 1 ] peripheral smears, bone marrow slides, fresh
capillary slides, and slides made for EDTA, heparinized, or oxalated bloods are
suitable to this staining technique. [ 2 ] If the SBB stain is old, increase the staining
time. [ 3]  Hematoxylin or Giemsa stains are satisfactory as a counterstain.

45      BRIEFLY DISCUSS THE PERIODIC ACID-SCHIFF REACTION AND DESCRIBE
 HOW IT IS PERFORMED.

Periodic acid-Schiff (PAS) stain detects the presence of intracellular glycogen.  This
stain detects muco-proteins, glycoproteins, and high molecular weight
polysaccharides.  A positive stain is a cell with a fuschia-pink color.  Staining
reactions are as follows:
[ 01 ]   Eosinophil granules do not take up the stin, but the cytoplasmic background
            stains positively.  This is a normal finding.
[ 02 ]   Early granulocytes show weak staining reactions, but the more mature forms react
            strongly.
[ 03 ]    Lymphocytes stain positive with varying degrees of intensity and patterns.  The
            staining reaction is usually weak.
[ 04 ]    Megakaryocytes and platelets stain positively with varying degrees of intensity
            and patterns. 
[ 05 ]   Monocytes stain positively with varying degrees of intensity and patterns.  The
            stain is usually a weak positive.
[ 06 ]   Nucleated red blood cells are negative, EXCEPT in patients with thalassemia
            and erythroleukemia (DiGuglielmo’s disease).
[ 07 ]   Basophils stain positive.

 The PAS reaction in abnormal hematological cells are as follows:
[A]  The erythroblast (in M6 erythroleukemia) stains positive.
[B]  The lymphoblast (in 80% of ALL cases) is positive.
[C]  The myeloblast (in 10% of the AML cases) is positive.

In  the PAS reaction, complex carbohydrates are oxidized to aldehydes to yield a red--
colored insoluble precipitate (aldehyde-fuscin-sulphurous acid molecule).  Basic fuscin
is responsible for the red color.

The procedure requires fixing the slides, followed by a rinsing sequence, after which the
slides are treated with periodic acid.  Stain in Schiff’s reagent, rinse, then counterstain
with hematoxylin.  Coverslip with permount (or its equivalent) and examine. 

Comments regarding this stain: [ 1 ] The mature neutrophils on the slide may be used
as positive controls. [ 2 ] If the patient is diagnosed with Burkitt’s leukemia, the
lymphoblasts will be PAS negative. [ 3 ] A Wright’s stained smear (even if years old)
may be PAS stained. [ 4 ] Schiff’s reagent is colorless or light yellow.  Add a drop to 37%
formalin, if a purple color, then the reagent is okay, otherwise discard.. If the Schiff’s reagent
turns pink, discard.   [ 5 ] PAS stain should not be used to try to differentiate types of
leukemia.  There is too much variability in the staining reactions to be reliable. [ 6 ] The
PAS stain is falling into disuse because of better specific cytochemistry marker
technology.

46      BRIEFLY DISCUSS THE LEUKOCYTE ALKALINE PHOSPHATASE STAIN
AND HOW IT IS PERFORMED.

 Synonyms: alkaline phosphatase score, alkaline phosphatase cytochemical test, and
alkaline phosphatase activity of neutrophils.  Leukocyte alkaline phosphatase (LAP)
enzyme is found in the cytoplasmic of mature neutrophils (neutrophil, band, and
meta--myelocyte) and this feature is used to differentiate a leukemoid reaction from
chronic myelogenous leukemia.  LAP activity increases as the granulocyte matures. 
The enzyme is found in the tertiary (microvesiuclar) granules of the neutrophil.  The
quantity of LAP enzyme varies within the neutrophil in various diseases.  LAP score
values range from a low of zero to a maximum of 400.  The normal value rage is 13 to
100.  Disorders with values less than 13 are chronic granulocytic leukemia, acute
granulocytic leukemia, marked eosinophilia, paroxysmal hemoglobinuria,
sideroblastic anemia,  hereditary hypophosphatasia, infectious mononucleosis, and
sickle cell anemia.  Values less than 13 have been reported in normal individuals. 
Elevated LAP scores may be observed in neutrophilic leukemoid reactions, last
trimester of pregnancy, corticosteroid therapy, multiple myeloma, polycythemia
vera, obstructive jaundice, myelofibrosis, and meningitis.

A blood specimen, fingersticks or heparinized specimens, is collected, smears
prepared and air dried.  The smear is fixed in an acetone and citrate buffer, then
stained in freshly prepared staining solution at a temperature between 18^oC and 26^oC. 
Smears may be counterstained in hematoxylin.  The patient’s smear should be stained
along with a negative/normal  and positive control to validate the staining
characteristics.  The blood from a woman in the 3rd trimester or a female on oral
contraceptives will provide a positive control and the blood from a healthy individual
will serve for a normal control.. 

To perform the LAP score, count 100 neutrophils and bands.  Grade the degree of
staining as follows:
[ 01 ]    Zero = no evidence of stain,
[ 02 ]   1+ = slight staining, very diffuse and faint without distinctive granular
                     appearance,
[ 03 ]   2+ = pale stain with small amount of granular appearance,
[ 04 ]   3+ = strong coloration, with moderate granular appearance,
[ 05 ]   4+ = intense coloration with large amount of granular, practically obscuring the
        cytoplasm. 
See page 401 - 402 of Rodak's Hematology (3rd Edition) for examples of LAP staining
with degrees of reactivity.

The test principle employs a substrate, such as naphthol AS-BI phosphate, which is
hydrolyzed in the presence of leukocyte alkaline phosphatase enzyme.  The
hydrolyzed substrate complexes with a dye that precipitates at the site of
enzyme activity.  When performing a LAP test, use slides with a monolayer,
so that the neutrophils do not touch the RBC’s.  If a thick slide is used, it is very
likely to falsely elevate the LAP score.  Do not attempt to include cells other than
neutrophils and bands in the count.  The slides once made, tend to deteriorate
quickly. Perform the LAP score evaluation quickly.  Do not allow the stained slides
to remain in direct light.   Control slides can be prepared and held ahead of time if
fixed and wrapped in a plastic film such as  parafilm.  Store such slides in a -70^oC
freezer.  If the lab uses commercial LAP staining kits, follow the manufacturer’s
directions closely.  Note normal value may vary from lab to lab.  Other normal value
reported are [ 1 ] 11 to 95 and [ 2 ] 30 to 185.

47        EXPLAIN HOW TO CALCULATE A LEUKOCYTE ALKALINE PHOSPHATE SCORE.

100 neutrophils and bands are counted and each cell is graded 0 to 4+.  Step one:
multiply the number of cells in each LAP grade times its rating.  Add the scores in the
five categories to determine the LAP score.  See the following example.

                        grade               # cells counted            value calculated
                           0                                30                                 0
                           1+                             35                                35
                           2+                             20                                40
                           3+                             10                                30
                           4+                               5                                20
                                                                        Total score       125

This test should be reported out as abnormal when using the normal range as
13 to 100. Negative/normal and positive controls should also be reported.

 48        BRIEFLY DESCRIBE THE ESTERASES OF GRANULOCYTES
AND MONOCYTES.

The esterases are a class of lysosomal enzymes that hydrolyze aliphatic and aromatic
esters at a neutral pH or less.  Esterases exist as isoenzymes and tend to be cell specific. 
Esterase isoenzymes 1, 2, 7, 8, and 9 are found in neutrophils.  These five isoenzymes
(designated as specific esterases only because they are found in neutrophils) may be
demonstrated with a naphthol AS-D chloroacetate substrate.  Those numbered 3, 4,
5, and 6 are characteristic to monocytes and other cells.  These four isoenzyme may
be demonstrated with α-naphthyl acetate substrate. Those designated as 2 and 4 type
isoenzymes may also be demonstrated with α-naphthyl butyrate substrate. Isoenzymes
3, 4, 5, and 6 have been arbitrarily designated as non-specific esterases.  The
characteristic chemical reactions of granulocytes and monocytes helps in the diagnosis
of granulocytic and monocytic leukemias.

 49        BRIEFLY DISCUSS CHLOROACETATE ESTERASE STAIN AND HOW
IT IS PERFORMED.

Synonym: Specific esterase, naphthol AS-D chloroacetate.  The myeloid leukocytes
contains a group of lysosomal enzymes designated as esterases.  Chloroacetate esterase
is a specific enzyme substrate that reacts with isoenzymes 1, 2, 7, 8, and 9; found in
granulocytes with myeloblasts and promyleocytes often demonstrating a positive stain. 
Auer rods stain positive.  Tissue mast cells stain positive, but monocytes and basophils
(of peripheral blood)  tend to demonstrate faint staining characteristics.  RBC’s,
lymphocytes, plasma cells, megakaryocytes, eosinophils, and nucleated red blood cells
do not contain this enzyme.  This staining technique is used to identify precursor cells
in acute myelogenous leukemia and aids in differentiating myelogenous cells from
monocytic cells.

Air dry the blood smear, then fix in the buffer solution, and next incubate in the
incubation mixture (containing naphthol AS-D chloroacetate) to release the naphthol
which combine with the dye in the incubation mixture and precipitates in the site of
enzyme activity.  The enzyme activity will show up as precipitate-like reddish-brown
colored granules.

50     BRIEFLY DISCUSS α-NAPHTHOL BUTYRATE STAIN AND HOW IT
IS PERFORMED.

Synonym: Non-specific esterase, α-naphthol butyrate.  This is an enzyme stain that is
used to help differentiate between granulocytic and monocytic leukemias.  The
α-naphthyl butyrate substrate is acted upon by the isoenzymes 2 and 4.  These two
isoenzymes produce a strong reaction in monocytes, a weak reaction in
megakaryocytes, and a focal reaction in lymphocytes. This stain is considered as not
staining granulocytes, lymphoblasts, plasma cells, or megakaryoblasts.  A positive
test is a pattern of staining characterized as a dark reddish precipitate in the
cytoplasm of the cells. 

A blood smear is prepared, air-dried, then fixed in a fixative buffer.  Next, incubate
the slide in an incubation mixture (containing α-naphthol butyrate).  Rinse and
counterstain with hematoxylin.  Use normal blood smears or bone marrow
preparations as controls.

51     BRIEFLY DISCUSS  α-NAPHTHOL ACETATE STAIN AND HOW IT
IS PERFORMED.

Synonym: Non-specific esterase, α-naphthol acetate.  This enzyme substrate
differentiates as does α-naphthol butyrate but is acted upon by isoenzymes 3, 4, 5,
and 6.   Monocytes and histiocytes stain strongly and megakaryocytes stain weakly. 
Lymphocytes demonstrate focal staining.  A positive test is a reddish-brown
precipitate in the cytoplasm of the cells. 

Prepare blood smear as described in objective 46.  Incubate the smear in an
incubation mixture containing α-naphthol acetate.  Rinse and counterstain with
hematoxylin.  Use normal blood smears and bone marrow preparations for controls.

 52      BRIEFLY DISCUSS NITROBLUE TETRAZOLIUM (NBT) STAINING
AND HOW IT IS PERFORMED

The nitroblue tetrazolium (NBT) stain is a “redox” dye that can be reduced to a purple
colored formazan compound by neutrophils.  The test has been used to differentiate
between bacterial and viral infections.  The procedure requires determining the
percentage and absolute number of neutrophils that reduce the colorless dye to a water-
soluble black fromazan deposit within the cytoplasm of neutrophils.  In a normal
individual, less than 10% of the neutrophils will be NBT-positive.  In a bacterial
infection, it is not unusual for 70% of the neutrophils to contain precipitated
formazan. 

The lab will collect heparinized blood.  A volume of blood is added to an equal volume
of NBT reagent.  The tube is mixed and incubated for 10 minutes at 37^oC.  A drop of
incubated mixture is transferred to a slide, and smear made.  The slide should be made
thicker than normal to minimize leukocyte damage.  The smear is air dried then
stained with Wright’s stain.   Perform a differential WBC on a separate smear stained
with Wright’s stain.  Count 100 intact neutrophils and determine what percentage
contains formazan.  Do NOT count damaged or distorted neutrophils.  Calculate the
absolute neutrophil count and multiply with the % of formazan WBC’s to obtain the
absolute NBT positive WBC count.  Buffy coat preparations may be used to increase
 the number on neutrophils on the slide.  The buffy coat preparation, carefully handled,
often demonstrates neutrophils with less distortion.  CAUTION:   Blood may be collected
with a plastic syringe or in heparinized capillary tubes.  If collecting blood in the plastic
syringe, handle the blood gently, and add one mL of blood to a tube with 20 units of heparin. 
Mix the blood by tilting and do not permit the blood to come in contact with the stopper of
the tube.  It is important to be sure that the ratio of blood to heparin is correct as an
excess of heparin may cause a false positive test. 

53      BRIEFLY DISCUSS TERMINAL DEOXYNUCLEOTIDYL TRANSFERASE
TEST AND HOW IT IS PERFORMED.

The terminal deoxynucleotidyl transferase (TdT) test is a means of identifying
lymphoblasts (primitive lymphoid cells).  This is an enzyme, deoxyribonucleic acid
polymerase, found in the nucleus of pre-B lymphocytes and T lymphoblasts.  This
enzyme identifies L1 and L2 acute leukemias.  This test can differentiate acute
lymphocytic leukemia from acute myelogenous leukemia.  There are three methods
used to assay TdT: immunofluorescence, immunoperoxidase technology, and
radioimmunoassay.  

Conduct the test as-soon-as-possible.  Do not stain if the slide is over seven days old. 
Fix the test slides, then rinse and hydrate in a phosphate buffer solution.  Apply
antibody to the slide and incubate for 30 minutes.  Rinse.  Apply the second antibody
and incubate for 30 minutes.  Rinse, dry, and prepare for examination.  The nuclei
of positive cells will fluorescence at 496 nm.  Record degree of fluorescence from zero
to 4+

54     BRIEFLY DISCUSS HEINZ BODY STAINING AND HOW IT IS PERFORMED.

The Heinz body (also called a Heinz-Ehrlich body) cannot be visualized in the 
erythrocyte stained with Wright’s stain.  They are formed from precipitated
hemoglobin, usually in size from 1.0 to 3.0 μM in diameter.  If they appear singly or
double, they tend to be large, but if there are several they will be smaller.  Heinz bodies
tend to lie close to the RBC membrane.  It is possible that they may be confused with
basophilic stippling if numerous and small. Basophilic stippling is a punctate
phenomenon due to the presence of aggregates of ribosomes.  The presence of Heinz
bodies are due to erythrocyte injury.  Drug poisoning and splenectomy result in the
appearance of Heinz bodies.   Supravital stains such as brilliant cresyl blue, crystal
violet, or methyl violet must be employed.  Staining is accomplished by mixing equal
volumes of blood and supravital stain, mixing, and incubating for 15 minutes. 
Prepare a blood film from the mixture and air dry.  Counterstaining is not required. 
Examine the slide for Heinz bodies under oil immersion.  To calculate the percentage
of Heinz bodies, the procedure used for determining the retic count will work.
            [ 1 ]        Count the # of Heinz bodies seen in 1000 RBC’s.
            [ 2 ]        Divide the number of RBC’s containing Heinz bodies
                           by 1,000.
            [ 3 ]        Multiply the value obtained in [ 2 ] times 100.
            [ 4 ]        Sample problem.   Assume that 65 RBC in 1,000 contained
                           Heinz bodies.

                                         65 RBC’s with Heinz bodies
            Heinz bodies  =   ----------------------------------- (×) 100
                                                1000 RBC's counted

            Heinz bodies = 0.065 (×) 100 = 6.5%

A score of 5% or greater is indicative of RBC damage due to toxic agents, either in
treatment regimes or accidental poisoning.  Other causes are Hemoglobin H disease,
G6PD deficiency, glutathione reductase deficiency, glutathione peroxidase deficiency,
triosephoshate isomerase deficiency, or splenectomy. 

55      BRIEFLY DESCRIBE THE ACID PHOSPHATASE STAIN FOR HAIRY
CELL LEUKEMIA.

Acid phosphatase is an enzyme located in all hemopoietic cells.  Acid phosphatase
exists in seven isoenzyme forms (0, 1, 2, 3, 3b, 4, and 5).  All of these isoenzymes will
enzymatically hydrolyze the naphthol AS-BI phosphoric acid substrate to yield
insoluble naphthol, which reacts with a chromogen to form a red colored azo dye. 
This stain has been used to help identify T-lymphocyte acute leukemia.  The hairy
cells of leukemic reticuloendotheliosis are abundant in isoenzyme 5.  If the other
isoenzyme enzymes (0, 1, 2, 3, 3b, and 4) are tested in the presence of L(+) tartaric
acid, no enzymatic activity is demonstrated.  The “hairy cell” will show a positive
reaction (varying shades of reddish color) due to the presence of isoenzyme 5.

56        BRIEFLY DESCRIBE BLOOD SMEAR PREPARATION FOR PARASITES.

Malaria is the most commonly studied parasite in blood.  There are four species of
malarial parasites.  Other blood parasites that may be encountered are Babesia
organisms, Trypanosoma species, and Leishmania species.   To prepare the blood, use
finger tip or fresh EDTA anti-coagulant blood.  Prepare 2 - 3 thin smears using the
wedge technique.  Allow to air dry.  To prepare the thick smears, place 2 - 3 drops of
blood in the center of the slide.  Use the corner of a second slide, spread the drop of
blood to the size of a dime and allow to air dry.  (NOTE: Dry for 12 hours, protected
from dust, in a petri dish set up.  Proceed as follows: [1] Fix the thin smears in
methanol, but DO NOT fix the thick smears. [2] Place the thin and thick blood smears
in 1/10 dilution of Giemsa stain, using buffered distilled water at pH = 6.8.  Allow to
stain from 30 to 60 minutes.  (NOTE: The Giemsa stain must contain Azure B stain. 
Azure A stain is not an effective parasite stain.) [3] Remove the slides from the
Giemsa stain and gently rinse under running tap water and allow to air dry.  View
microscopically.  On the thick smear, WBC nuclei and platelet debris will be seen.

 57     DISCUSS THE BLOOD SMEAR STRATEGIES AND AVOIDANCE OF
ARTIFACTS AND ERRORS THAT MAY BE OBSERVED IN A BLOOD SMEAR.

The following should be adhered to minimize the appearance of artifacts:
[ 01 ]        Do not stain a peripheral blood smear until it is properly fixed.  Use methanol
                fixative adjusted to a pH = 8.4. 
[ 02 ]        Watch the staining time.  Do not allow Wright’s stain to remain on slide or
                 methanol will evaporate and cause precipitation of dye molecules.
[ 03 ]        If water is present in methanol, ring-shaped, refractive artifacts will appear
                on erythrocytes.  Do not confuse with RBC inclusions.
[ 04 ]        Do not evaluate RBC hemoglobin content at end of slide.
[ 05 ]        Do not evaluate smear along edges, the cells tend to be distorted or elongated.
                This is an artifact of spreading.
[06 ]        When examining the thin edges of the film and crenated RBC’s (or
                echinocytes) are noted, if the spicules are uniform, do not report, this is an
                artifact.
[ 07 ]        If you wipe the oil from a stained smear, it is possible for the tissue to
                damage RBC’s, causing them to appear as schistocytes.  Do not wipe! 
                Dab the oil from the slide. 
[ 08 ]        If you see a phenomenon of some type and it appears to be in straight line,
                 it is probably an artifact.
[ 09 ]        When you are viewing a slide before performing a differential and you
                 observe target cells in one area, but not others, do not report.  If the target
                 cells are distributed randomly across the slide, then report as 1+, 2+, 3+ or 4+.
[ 10 ]       If you see a RBC with a distinct colored outer circle with a well defined clear
               center, without gradation, it is an artifact, not hypochromia.  Hypochromia
               is characterized by gradation from the outer edge to the central area of pallor.
[ 11 ]      Wright’s stained slides will fade over time unless mounted with a cover glass.
[ 12 ]      Slides with dirt/grit may result in precipitation of stain.

Other “things” that can cause RBC artifacts are [A] delays in making smears, [B] too
hot or too cold temperatures in lab, [C] the smear dries to slowly, [D] polycythemia
(increased blood viscosity), [E] the presence of abnormal proteins, [F] pH to acidic or
alkaline, causing changes in the erythrocyte’s internal environment, [G] pressing too
hard on the spreader slide as the smear forms.

58        DESCRIBE CRITERIA THAT IS HELPFUL IN EVALUATING A BLOOD SMEAR

[ 01 ]   Perform the differential count in the monolayer in the middle portion of the slide.
[ 02 ]    Avoid the thick regions because the cells tend to “bunch up” and obscure
             abnormalities.  If the entire slide appears to have cells stacked on top of each
             other, the slide is too thick.
             A.    When the smear was made, the angle of the spreader slide was to large.
             B.     To correct, make a new slide with a smaller angle of the spreader slide.
[ 03 ]    If the end of the smear does not have a feather edge, the angle of the spreader
             slide was too large or its edge had cracks and/or chips.  Use a spreader slide
             with a smooth, sharp edge.
[ 04 ]    If the cells are to faint to be seen, the staining time was too short.  Make a new
            smear and repeat by staining a longer time.
[ 05 ]    If the problem in #4 is not attributed to inadequate staining, look at the rinse
            buffer.  Its pH may be to acidic.  Resolve the problem by using an appropriate
            with a pH of 6.8.
[ 06 ]    If there are holes in the smear, the slide contained some form of Contamination
            on its surface.  Use only high quality slides to eliminate this problem.
[ 07 ]    If unidentifiable “things” are seen on the slide, it may be the result of using a
            dirty slide.
[ 08 ]    If the cells are so dark the nuclei cannot be distinguished, the slide was over
            stained.  Slides like these may have visible precipitate stain.  Make a new
            smear and stain a shorter time.
[ 09 ]    If the problem in #7 is not over staining, then the pH of the buffer rinse may be
            too alkaline.  Resolve the problem by using an appropriate buffer with a pH
            of 6.8.
[ 10 ]  Rate the staining qualities.
           A.    The thicker areas stain darker and have more artifacts.
           B.    Is the stain quality good or poor.  The slide may need to be restained.
[ 11 ]  The normal size of the RBC is 6.0 to 9.0 μm, with the 0 = 7.8 μM.
[ 12 ]  When evaluating the erythrocytes for abnormalities (such as cell size and shape, 
           it is recommended that the degree (if any) of anisocytosis, poikilocytosis, and
           hypochromia be noted.
           A.    If the RBC’s are distorted and proper classification is compromised, then
                    the anticoagulant being used may be a problem or the slide was not
                    allowed to dry properly.
           B.    Change to a different anticoagulant and/or allow the slide to thoroughly
                    air dry.
           C.    Making blood smears from a finger stick may be the best solution to
                    cell distortion.
[ 13 ]    If one or more of the three abnormalities are present, look for RBC inclusions
            examples: Howell-Jolly bodies, Pappenheimer bodies, and Heinz bodies).
[ 14 ]    When observing leukocytes, note the size of the cell.
          A.     Small is no smaller than an erythrocyte.  Lymphocytes are the only WBC’s
                    that are expected to fall in this size category.  This size ranges from 8.0 to
                    10.0 μm.   (Hint: The nucleus of a small lymphocyte is about the same size as
                     that of a RBC.)
          B.    Medium size describes most neutrophils.  This size ranges from 9.00 to
                  15.0 μM.
          C.    Large is characteristic of monocytes.  This size ranges from 14.0 to
                  20.0 μM.
[ 15 ]    Notice the shape of the nucleus.
          A.     Neutrophils have a segmented nucleus.
          B.      Bands tend to have a “U” or “C” shaped nucleus, but can be "S” shaped.
          C.     The nucleus of the lymphocyte tends to be round.
          D.     The monocyte is characterized by a convoluted, sprawling nucleus.
          E.      A notched nucleus may be observed in the lymphocyte and metamyelocyte.
[ 16 ]    Evaluate the texture of the nucleus.
         A.     If the nucleus is dense and dark, it is pycnotic (neutrophil).
         B.     A close knit nucleus is seen in the lymphocyte.
         C.     A ropy, spongy-like nucleus is typical for the monocyte.
         D.     Fine featured, with little or not texture, may indicate an immature
                   leukocyte.
[ 17 ]   Consider the chromatin pattern.
         A.     Is it smooth or coarse.
         B.     Is the parachromatin (light staining areas) visible ?
[ 18 ]      Are nucleoli present or absent?
[ 19 ]      Look at the cytoplasm of the leukocyte.
         A.     Large, distinctively colored granules feature the eosinophil and basophil.
         B.     The neutrophil has fine granules evenly distributed in the cytoplasm.
         C.     The lymphocyte has a homogenous, light blue colored cytoplasm.      
         D.     A grayish coloration characterizes the monocyte.
         E.     Compare the staining characteristics around the inside periphery of  the
                  cytoplasmic membrane with that on the outside of the nuclear membrane.
                  Very immature cells tend to exhibit basophilia at the periphery. 
                  Lymphocytes are characterized by light staining about the nucleus
                  (perinuclear halo).
         F.     Compare the ratio of the cytoplasm to the nucleus.

 [ 20 ]  Abnormalities to watch for in the leukocytes are:
         A.    Unusual granulation (example: toxic granulation in neutrophils due to a
                  severe infection).
        B.    Cytoplasmic vacuolation which may be observed in all WBC’s.
                a.    Vacuolated nutrophils observed in severe infections.
                b.    Infectious mononucleosis causes vacuolation in lymphphocytes.
                c.    Normal monocytes may exhibit some vacuolation.
                d.    Toxic chemicals/drugs can cause vacuolation in any WBC.
         C.    Disintegrating cells occur when the cytoplasmic membrane ruptures and the
                cytoplasm and nuclear contents are somewhat intact (the cell can still be
                 identified).  This occurs among all cells and is usually negligible.  If there
                 are large numbers present on the slide, that may indicate a pathology. 
                 Remember that such cells may be the result of making a slide from old blood
                 or improperly making a blood smear.  Follow lab protocol in reporting
                 disintegrating cells.  Some labs do not report the presence of a few
                 disintegrating  cells.
          D.   Smudge cells.  These are not disintegrating cells.  This cell is characterized by
                  the presence of oddly shaped nucleus and little evidence if any of cytoplasmic
                  material.  An occasional smudge cell is not significant.  Numerous smudge
                  cells may indicate a toxic or leukemic condition.
          E.   Inclusion bodies in the cytoplasm of leukocytes may be an indicator of a
                 pathological condition.   An example is the Dohle body, a distinct blue mass
                 in the cytoplasm of a neutrophil.
[ 21 ]    If the differential count is too high for certain leukocytes, one may counting the
           same fields more than once.  Repeat count and watch the scanning technique to
          avoid repeating fields.
          A.    If the problem can be related to differentiating the cell types, repeat the count
                  with a different technologist.

59        BRIEFLY DESCRIBE THE REPORTING STRATEGY FOR RBC’s.

It is generally acceptable to report the abnormalities as either present or absent.  If one
or two abnormalities are observed in the entire slide, do not report them.  Occasional
abnormalities may be seen in blood smears for normal, healthy individuals.  If the
laboratory requires that abnormalities be reported in degrees, the following is an
acceptable “rule-of-thumb”: slight = <15% of the field contains abnormalities,
moderate = approximately 20 to 59% of the field contains abnormalities, and
anything over 60% should be reported as marked.  If RBC inclusions are observed,
report as the number per 100 RBC’s.  The lab manual contain a descriptor sheet
with a reporting strategy for RBC’s.

60        BRIEFLY EXPLAIN THE SCHILLING HEMOGRAM/CLASSIFICATION.

Schilling (German pathologist) noticed that the granulocyte series increased in the
number of immature cells during pathological disorders.  He modified the Arneth
count to a simpler form to include the granulocytic evaluation.  The Schilling
hemogram is a WBC differentiation scheme that evaluates the percentage of neutrophils
per 100 WBC’s.  Schilling introduced the phrases “shift-to-the-left” to indicate more
immature granulocyte cells and “shift-to-the-right” to indicate an increase in mature
granulocytes.  His scheme was to set up the reading scale so that the more immature
granulocytes would be listed on the left and the mature forms on the right.  The scale
reads thus: [a] myeloblasts and promyelocytes, [b] myelocytes, [c] metamyelocytes-
slightly indented forms, [d] metamyelocytes-band form, and [e] segmented neutrophils. 
He determined the normal value to be: [a] myeloblasts, promyelocytes, and
myelocytes = 0%, [b] metamyelocytes-young form = 0%, [c] metamyelocytes-band
form = 1 to 5%, [d] neutrophils = 30 to 70%.  Schilling also identified two types of
shifts-to-the-left:
[1]    regenerative shift-to-the-left, characterized by a rapid rate of
         production of WBC’s with a significantly elevated WBC count. 
         He noted this shift in appendicitis and acute sepsis.
[2]    degenerative shift-to-the-left, characterized by lower WBC count
         and the number of immature granulocytes expected in circulation
         are depressed by toxins that interfere with the maturation of the
         granulocyte.  He observed this shift in typhoid fever, brucellosis,
         pernicious anemia, and TB.

Schilling pointed out that in the normal recovery of a patient, a shift-to-the-right would
occur characterized by an increase in lymphocytes and eosinophils before other clinical
symptoms became obvious.  If the shift-to-the-left persisted, it was a poor prognostic sign.

Schilling used this concept in performing the WBC differential, which was known as the
 Schilling hemogram.  It forms the basis for the way WBC differentials are performed
 today.  NOTE: If the WBC count is greater than 35,000/μL, count 200 WBC’s in
the “diff” instead of 100.

61        BRIEFLY EXPLAIN THE ARNETH COUNT. 

Arneth (German pathologist) classified neutrophils according to their age, based upon
the number of lobes in the nucleus of a neutrophil.  He described five age neutrophil
age groups:
   [ 01 ]    A single round or indented nucleus as the youngest cell = 5%,
   [ 02 ]    two distinct nuclear divisions as the next youngest cell = 35%,
   [ 03 ]    three distinct nuclear division as middle age = 41%,
   [ 04 ]    four distinct nuclear divisions = 17%,
   [ 05 ]    five or more distinct nuclear divisions as the oldest cell = 2%. 
This classification has some merit but is time consuming for routine laboratory use,
therefore it is seldom referred to.  It has used in determining hypersegmentation of
neutrophils in vitamin B₁₂ deficiency anemias.   Comment: Some hematologists consider
a five-lobed neutrophil to be hyper-segmented.

62      BRIEFLY EXPLAIN THE PRINCIPLE OF THE FILAMENT VS. THE
NON-FILAMENT CLASSIFICATION SCHEME. 

This was a classification scheme that stated that a filament cell included those cells that
contained a lobe or segment connected by a filament.  There were the neutrophils,
eosinophils and basophils.  The non-filament cells included myelocytes,
metamyelocytes, bands, lymphocytes, and monocytes.  It could also contain the more
immature forms not listed.  If this scheme were employed, a normal differential would appear as follows:
                                                Schilling-type          filamented/non-
non-filamented      normal range          differential             filamented diff.
    myelocytes             0%
    metamyelocytes      0%
    bands                   2 - 5%                   2
    lymphocytes        20 - 35%                30                              36%
    monocytes            2 - 6%                    4
filamented
    neutrophils          35 - 65%                 60
    eosinophils           1 - 3%                     3                              64%
    basophils              0 - 1%                     1

63      EXPLAIN HOW TO EVALUATE OR ESTIMATE THE NUMBER OF
PLATELETS AND LEUKOCYTES ON A BLOOD SMEAR.

Platelet evaluations are a routine part of the WBC differential.  The normal procedure
is to count 15 - 20 RBC’s and note the number of platelets present.  Normal is one
platelet per 15-20 RBC’s. If there is <1 platelet/15 - 20 RBC’s, the platelets are
decreased and the count is expected to be decreased.  If there are >1 platelet/15 - 20
RBC’s, the platelets are increased and the count is expected to be increased.  One
recommended procedure is to count the number of platelets in ten “oil-immersion
fields” and calculate an average number of platelets per “oif”.  The count is to be
conducted in the monolayer of the blood smear where the RBC's are not over--
lapping.  Next multiply the average number of platelets times 20,000 and this will give
ctimated  platelets/μL.  When you are estimating the number of platelets on a stained
blood film, report as average number of platelets per oil immersion field (oif).

If approximately 3 to 4 WBC’s are observed per “oif”, then the WBC count is
expected to be in the normal range.  If less than this, a low count.  If greater than
five WBC’s, the count is expected to be elevate

                                                            T A B L E
0 number of WBC’s         approximate WBC           approximate
 or platelets/ “oif”                count/μL              platelet count/μL
       1 - 4                              2,000 - 8,000                  30,000 - 60,000
         4 - 6                             8,000 - 12,000                   60,000 - 90,000
         6 - 10                          12,000 - 20,000                 90,000 - 150,000
        10 - 20                         20,000 - 40,000               120,000- 300,000

 64        LIST PROCEDURES IN WHICH MANUEL COUNTS STILL MAY BE
PERFORMED.

[ 1 ] Elevated leukemic WBC counts. [ 2 ] Platelet counts. [ 3 ] Spinal fluid.
[ 4 ] Synovial fluid. [ 5 ] When the automated cell analyzer breaks down.

65      EXPLAIN HOW TO CORRECT A WBC COUNT WHEN NUCLEATED RED BLOOD CELLS ARE PRESENT.

This correction is initiated when nucleated red blood cells (NRBC) are encountered
on a differential.  The number of NRBC’s must be enumerated per 100 leukocytes. 
A corrected count may be reported by using the following formula:

                                        # of uncorrected WBC’s (X) 100
corrected WBC count =  ---------------------------------------------------
                                       100 + Number of NRBC’s/100 WBC’s

Sample problem: [1] 25 NRBC’s counted on differential/100 WBC’s,
[2] uncorrected WBC count = 13,500 μL
                                         13,500 (x) 100               1,350,000
[3]  corrected WBC count =  ---------------------   =   ---------------  =
                                         100 (+) 25                       125
[4]  answer to the  problem is:   10,800/μL

66    DESCRIBE THE HEMOCYTOMETER.

The hematocytometer is a counting chamber with identically duplicate ruled counting
areas on a raised platform.  A cover glass is required for placement over the platforms
to provide a depth of 0.1 mm.  Refer to the illustration below as you read through this
paragraph. Each ruled area is a 3.0 mm square divided into nine equal size square of
1.0 mm on each side.  The area of the large square is 9.0 mm² and the area of each of
the nine small squares is 1.0 mm².  The volume of the large square is obtained by
multiplying the area (9.0 mm²) times the depth (0.1 mm) which equals 0.9 mm³.  The
volume of each of the 1.0 mm squares is 0.1 mm³.   The four corner squares are divided
into 16 smaller squares each.  Each of these smaller squares measures 0.25 mm on each
side.  The area of each of the smaller squares is 0.0625 mm² and the volume equals
0.00625 mm³.  The outer large four corner squares have been used for counting
leukocytes.  The center large square is subdivided into 25 smaller squares.  Each of the
25 subdivided squares are divided into 16 still smaller squares.   The dimensions of each
of the 25 subdivided squares are 0.2 mm on each side.  The area of one of these squares
is 0.04  mm² and the volume would be 0.004 mm³.   The dimensions of each of the
sixteen smaller squares is 0.05 mm on each side.  The area of each of these tiny squares
becomes 0.0025 mm and the volume is 0.00025 mm³.  (NOTE:  μL and cu.mm and  mm³
are equivalent measures.)   All hemocytometers are identical in meeting the area and
volume specifications.  There may be some variation is the way a hemocytometer
may appear or in the number of lines to make the rulings.

67        DISTINGUISH BETWEEN AN ABSOLUTE AND RELATIVE COUNT.

The absolute count means to be free from mixture, to have no restrictions.  In the
laboratory, it is an expression of the numbers of each cell type/μL of blood.  It is a
means of imparting additional information.  It is a mathematical calculation that
 determines the actual number of a cell type so that its increase or decrease may be
known.  The calculate the absolute count, the relative count must be known.  Use
the following formula:

absolute count =   total WBC count (x)  relative count
   (WBC/μL)                 (WBC/μL)        % WBC/μL)

Table of Absolute and Relative Values (Normal WBC Range)

                                          relative                          absolute count
                                           count (%)                            (cells/μL)
          Total WBC’s                                                      5000 - 10,000
          Myelocytes                       0                                        0
          Metamyelocytes              0 - 1                                  0 - 100
          Bands                             2 - 5                                100 - 500
          Neutrophils                   35 - 65                          1750 - 6500
          Eosinophils                      1 - 3                             50 - 300
          Basophils                         0 - 1                                0 - 100
          Lymphocytes                  20 - 35                          1000 - 3500 
          Monocytes                       1 - 6                               100 - 600

Example #1.     If the WBC count = 25,000/μL and the relative lymphocyte
                      count =  76%, then the absolute count would be 18,750/μL.

Example #2.     If the WBC count = 7,300/μL and the relative neutrophil
count = 60%,   the absolute count would be 4,380/μL.

68        DESCRIBE THE THOMA CELL COUNTING PIPETTE.

 The Thoma cell counting pipette is a calibrated glass pipet with a bulb for a diluting
chamber.  There are two types of pipets, one for WBC counting (characterized by a
clear or white mixing button in the mixing chamber) and the second for RBC counting
(identified by the red mixing button in the diluting chamber).  Each has a pipet stem
with calibration marks.  Both pipets do not measure in mLs, but in parts.  Each pipet
is designed to give a specific dilution. 

The WBC pipet can dilute from 1:10 to 1:100.  Most WBC pipets contain ten
calibration marks designated as 0.1 to 1.0.  A final calibration mark is located on the
opposite side of the bulb (designated by 11).  The volume in the stem is 10 times less
than that of the bulb.  Blood, pipetted to the 0.5 mark, then diluted to the 11 mark
provides a 1:20 dilution. Note the dilutions possible using the WBC pipet and
pipetting blood (or any body fluid) to the to the 0.1 mark and diluted to the 11
mark give a 1:100 dilution.  

The red blood cell pipet is calibrated with similar marks, but with one difference, the
11 mark becomes a 101 mark.  The volume of the stem is 100 times less than that of the
 bulb.  The dilutions possible with the RBC pipet are increased ten fold.  Blood (or any
body fluid) drawn to the 0.5 mark and diluted to the 101 mark yields a 1:200 dilution. 
Pipetting to the 1.0 mark yields a 1:100 dilution.

                                                  T A B L E
A body fluid drawn the the designated mark on the stem and diluted to the 11 or 101
mark on the opposite side of the bulb gives the following dilutions.
         Mark on the stem            WBC pipet dilution        RBC pipet dilution
                    0.1                                1:100                            1:1000
                    0.2                                1:50                              1:500
                    0.3                                1:33                              1:333
                    0.4                                1:25                              1:250
                    0.5                                1:20                              1:200
                    0.6                                1:17                               1:167
                    0.7                                1:14                               1:142
                    0.8                                1:12                               1:125
                    0.9                                1:11                               1:111
                    1.0                                1:10                               1:100  

69        DESCRIBE A TRENNER PIPET.

The Trenner pipet differs from the Thoma pipet in the way the stem is joined to the
mixing bulb.  The stem inserts into the bulb so that the end is flat, polished, and
at right zangles to the longitudinal axis.  The means that blood can be drawn into
the stem by capillary action and will fill the stem, automatically stopping at the end
 of the stem.  Each Trenner pipet is calibrated to dilute to a designated volume.
           

70      IDENTIFY THE SOURCES OF ERRORS IN THE HEMOCYTOMETER
COUNTING PROCEDURE.

Equipment related errors include:
[1]        moisture in the counting chamber,
[2]        moisture in the diluting pipet,
[3]        dirty glassware. 
Blood sampling related errors include:
[1]        a capillary puncture that does not flow freely,
[2]        clots present in the blood or other body fluid,
[3]        incorrect ratio of blood and anticoagulant,
[4]        incorrect ratio of blood (or body fluid) to diluent. 
Test performance errors include:
[1]    incorrectly filling the pipet,
[2]    failure to adequately mix the diluted sample in the pipet,
[3]    failure to discard the first three drops of diluted sample to clear the stem,
[4]    failure to fill the hemocytometer,
[5]    allowing bubbles to be trapped in the hemocytometer,
[6]    error in enumerating the cells in the counting area,
[7]    calculation errors. 
Inherent errors include:
[1]   the distribution of cells in the chamber (if the distribution is uneven, the error f
        actor increases)
[2]    counting too small a population of cells (the greater the number counted, the less
         the error).  Note: There is an inherent error variation of 4.5% in the
         hemocytometer. 

71        DISCUSS THE IMPORTANCE OF THE WBC COUNT.

Leukocyte numbers fluctuate in health and disease.  If the WBC count drops below
the normal range (5000 to 10,000/μL), then the condition is leukopenia.  If the count
is elevated over 10,000/μL, then the condition is leukocytosis.  These conditions are
due to depression or stimulation of bone marrow and other elements.  The WBC
count in children tends to fluctuate more widely than adults in disease.  White cell
counts tend to be higher in the afternoon than in the morning.  Strenuous exercise and
emotional ups-and-downs will promote an increase in the WBC count.  WBC counts
can be employed to follow the effectiveness of treatment therapies.

Causes of leukopenia are [A] measles, [B] hepatitis, [C] systemic lupus erythematosus,
[D] radiation treatments, [E] rheumatoid arthritis, [F] influenza, [G] cirrhosis of the
liver, [H] antibiotic therapy, [I] hormone therapy, [J] gram negative septicemia,
[K] hemodialysis, [L] typhoid fever, [M] brucellosis, and [N] chemotherapy. 

Causes of leukocytosis are: [A] appendicitis, [B] pneumonia, [C] leukemia,
[D] meningitis, [E] abscesses, [F] uremia, [G] pregnancy, [H] ulcers,
[I] rheumatic fever, [J] chicken pox, [K] parasite infestations, [L] burns,
[M] stress, and [N] allergies. 

72        DISCUSS THE IMPORTANCE OF THE RED BLOOD CELL COUNT.

Red blood cell counts generally contribute little clinical information.  Hemoglobin and
hematocrit determinations are usually preferred.  The RBC count is important for the
calculation of the indices.  Erythrocyte numbers fluctuate in both health and disease. 
A decrease in the RBC count is known as erythropenia or oligocythemia.  An increase
in the RBC is known as erythrocytosis.  A decrease or increase in the RBC count will be
due to a depression or stimulation of the bone marrow elements.

 Erythropenia may be caused by [A] a wide variety of anemias, [B] lead poisoning,
[C] pre-leukemic states, [D] infections, [E] maturation disorders, [F] abnormal
intravascular hemolysis, [G] bone marrow aplasia, and [H] hemorrhage. 
Erythrocytosis may be caused by [A] dehydration, [B] stress, [C] polycythemia vera
rubra, [D] bone marrow hyperplasia, [E] benign polycythemia, and [F] erythroleukemia.
 

73    THE IMPORTANCE OF MONITORING FOR ERRORS.

The Clinical laboratory is concerned about quality and accuracy of the tests that
are reported to primary care givers.  The laboratory monitors where these errors
can appear that will affect the accuracy of test results.   These errors can occur
prior to the test analysis and if they manifest, they are called preanalytical errors
or variables.  If the error occurs during the testing process, then it become an
analytical error.  If the error appears after the test is performed and reported, then
it is known as a post-analytical error.

The preanalytical error occurs before the test is performed.   This error source can
occur at the beginning of test ordering and flling out the requisition.  Examples of
this type of error includes:
01    duplicate or missing requisitions
02    tests omitted from the requisition
03    incorrect ordering of tests
04    patient identification error
05    incorrect blood collection
06    specimen transport error
07    specimen handling/processing in the lab

Analytical errors occur during the testing process.  Examples of these errors are:
01    deteriorated or wrong reagents
02    any instrument malfunction
03    laboratorian error
04    incorrect recording of test results

When the lab determines that the testing process was conducted in a flawless
manner and there were no mistakes, the report is ready to be released.  At this
point in time, any errors that take place are postanalytical.   Examples of these
are:
01    failure to notify the physician of critical values.
02    failure to report test results in a timely manner.
03    placement of report in the chart of the wrong patient 
04    miscommunications that are detrimental to the patient regarding the tests
        performed. 
 

This web site is maintained by Whitney Williams, wwilliam@astate.edu

This page last updated 07/28/08